Significance

FIC-domain enzymes are found in all kingdoms of life and catalyze posttranslational modifications of various target proteins to modulate their function. Because the vast majority of Fic proteins are expressed in an inhibited form, their physiological importance has escaped attention for a long time. This article reveals an autonomous mechanism of inhibition relief for class III Fic proteins, which hinges on autoadenylylation of an inhibitory helix. Because the process occurs in cis, the Fic enzyme constitutes a molecular timer that operates independent of enzyme concentration. Furthermore, we show that Fic-mediated adenylylation of DNA gyrase leads to bacterial growth arrest. Thus, the time-dependent inactivation of DNA gyrase may serve as a switch to bacterial dormancy under starvation or other stress conditions.

Abstract

Filamentation induced by cyclic AMP (FIC)-domain enzymes catalyze adenylylation or other posttranslational modifications of target proteins to control their function. Recently, we have shown that Fic enzymes are autoinhibited by an α-helix (αinh) that partly obstructs the active site. For the single-domain class III Fic proteins, the αinh is located at the C terminus and its deletion relieves autoinhibition. However, it has remained unclear how activation occurs naturally. Here, we show by structural, biophysical, and enzymatic analyses combined with in vivo data that the class III Fic protein NmFic from Neisseria meningitidis gets autoadenylylated in cis, thereby autonomously relieving autoinhibition and thus allowing subsequent adenylylation of its target, the DNA gyrase subunit GyrB. Furthermore, we show that NmFic activation is antagonized by tetramerization. The combination of autoadenylylation and tetramerization results in nonmonotonic concentration dependence of NmFic activity and a pronounced lag phase in the progress of target adenylylation. Bioinformatic analyses indicate that this elaborate dual-control mechanism is conserved throughout class III Fic proteins.

Fic (filamentation induced by cyclic AMP) proteins containing the FIC domain (pfam 02661) are found in all kingdoms of life. FIC domains encode enzymatic activities that modulate target protein function by diverse posttranslational modifications (1, 2). The vast majority of known Fic proteins are AMP-transferases that use ATP to catalyze the transfer of an AMP moiety onto a target hydroxyl side chain (3, 4). This reaction is akin to the situation in protein kinases, which catalyze γ-phosphate transfer onto target side chains.

Only a few Fic targets have been identified to date (2). IbpA (4) and VopS (3), two bacterial FIC-domain effectors that get translocated into host cells, catalyze the adenylylation of Rho GTPases, resulting in cytoskeleton collapse. Most recently, we have found that a subset of bacterial Fic proteins adenylylates DNA gyrase and topoisomerase IV, which leads to their inactivation and cellular growth arrest (5). The structure of a FIC-domain/target complex (6) and the catalytic mechanism have been determined (6, 7). However, the biological functions, as well as the molecular mechanism, of the vast majority of Fic proteins have remained elusive. Intriguingly, in vitro automodification has been demonstrated for most Fic proteins that have been described so far (6, 8⇓⇓⇓⇓⇓⇓–15), but its physiological relevance has remained unclear.

Recently, we have shown that Fic-mediated adenylylation is tightly regulated (8). In the native state, a helix partly obstructs the ATP binding site with a strictly conserved Glu blocking ATP γ-phosphate binding, thereby preventing productive/competent substrate binding. The inhibitory α-helix (αinh) is either located on a separate protein that forms a tight toxin/antitoxin complex with the Fic enzyme (16) or at the N- or C-terminal position within the same polypeptide chain. These three possibilities lead to a classification of Fic proteins into classes I, II, and III, respectively (8). Mutation of the inhibitory Glu to Gly relieves autoinhibition, thus boosting both target and autoadenylylation (8, 9). However, the identity of the intrinsic or extrinsic factors that in vivo expulse the inhibitory Glu of αinh, and thereby relieve autoinhibition, has not been investigated.

Fic proteins have evolved in bacteria and have spread by horizontal gene transfer into all domains of life (17). Despite structural conservation of the basic FIC-domain fold, there is significant sequence diversity among class I and II Fic proteins. Additionally, these two classes are frequently combined with other protein domains toward multidomain arrangements, demonstrating a high functional plasticity and adaptability (2). In contrast, class III Fic proteins are highly conserved single-domain proteins even though they are found scattered across all classes of Proteobacteria (8). This conservation suggests that class III Fic proteins are stand-alone autoregulated functional entities that act in a plug-and-play manner upon acquisition by horizontal gene transfer.

Here, we dissect the regulatory mechanism of NmFic from Neisseria meningitidis as representative for the class III Fic proteins. First, we identify the B-subunit of DNA gyrase as the main bacterial target of NmFic, as for class I Fic proteins (5). We then reveal the crucial role of autoadenylylation in the activation of class III Fic proteins and the opposing role of oligomerization, resulting in a peculiar and intriguing NmFic concentration dependence of target adenylylation. Ultimately, because the oligomerization interfaces are either highly conserved or covaried, and because the modifiable residue Y183 is strictly conserved in class III Fic proteins, we anticipate that the combination of oligomer dissociation and subsequent cis-autoadenylylation is the major regulatory mechanism of class III Fic proteins.

Results

NmFic Adenylylates DNA Gyrase.

We have previously shown that expression of inhibition-relieved mutants of various bacterial Fic proteins slowed down the growth of ectopically expressing Escherichia coli (8). This growth reduction correlated with in vitro adenylylation of an endogenous protein with a mass of about 90 kDa. Subsequently, it was shown that the class I Fic protein VbhT adenylylates the B-subunit of the bacterial topoisomerases DNA gyrase (GyrB) and topoisomerase IV (ParE) (5). The modification affects a conserved Tyr of the ATP binding-site lid, thus interfering with ATPase and topoisomerase activity.

Because expression of (class III) NmFic in its inhibition-relieved form (NmFicE186G) also slows down E. coli growth (8), we tested whether NmFic modifies the same targets (Fig. 1). Corroborating our previous findings (8), NmFicwt does not adenylylate any of the target proteins. In contrast, NmFicE186G efficiently adenylylates N. meningitidis GyrB and the orthologous E. coli and Mycobacterium tuberculosis proteins, but not ParE of N. meningitidis or E. coli. Mutation of the E. coli GyrB acceptor site (Y109F or Y109A variant) completely abrogates the effect (Fig. 1), confirming the predicted modification. In addition, independent of the presence of the target, strong autoadenylylation of NmFicwt and NmFicE186G is observed (Fig. 1), which is investigated further below.

Conserved Tetrameric Organization of Class III Fic Proteins.

NmFic invariably shows a tetrameric arrangement (Fig. 2 and Fig. S1) in different crystal forms and ligation states, even when the C-terminal helix is absent or disordered (Fig. S1 A–C). The tetramer is of 222 symmetry (dimer of dimers), and its formation involves two independent interfaces (Fig. 2 A–C and Movie S1). Interface 1 is mainly mediated by the apolar interactions of F70 and Y77, with their respective symmetry mates, and by R71 and E102, which form two isologous salt bridges (Fig. 2B). The interface 1 residues are strongly conserved among the 197 analyzed sequences of class III Fic proteins. Similarly, interface 2 is formed by apolar residues L155 and F159, as well as the two isologous salt bridges of R149 and E156 (Fig. 2C). These residues are not strongly conserved, but the interacting residues show strong covariation with the striking charge reversal of the E-R to K-E salt bridges in a subset of the class III proteins. Taken together, conservation and covariance of the surface residues of NmFic suggest that tetramer formation is of functional importance for class III Fic proteins.

Structural and oligomeric analysis of NmFic. (A) Cartoon of the NmFic structure (PDB ID code 3S6A) with the active site motif highlighted in yellow and the target binding site (flap) in blue. Important residues are shown in full with side-chain carbon atoms of active site residues colored in yellow, modifiable Tyr in purple, and charged residues mediating oligomerization in magenta (interface 1) and green (interface 2). The αinh is shown in olive-green with the inhibitory Glu (E186) shown in full. (B) View of the crystal structure of the NmFic tetramer (222 symmetry with the twofold axes indicated) with the subunits distinguished by color (Left) and close-up views of the dimerization interface 1, with a contact area of 835 Å2 (Right). Side-chain carbon atoms are colored according to the conservation score [ConSurf (28)]. (C) Ninety-degree rotation of the crystal structure shown in B (Left) and close-up view of the dimerization interface 2, with a contact area of 425 Å2 (Right). Side-chain carbon atoms are colored according to covariance probability [Gremlin (29)]. In B and C, asterisks denote residues from the neighboring molecule. (D) Dynamic oligomerization equilibrium of NmFicwt and variants. (Top) Concentration dependence of the apparent molecular weight as determined by SEC-MALLS for NmFicwt and oligomerization interface variants. (Bottom) Tetrameric fraction xQ of NmFicwt in the absence and presence of 5 mM ATP and autoadenylylated NmFicwt-AMP in the presence of 5 mM ATP. The lines represent the nonlinear least squares fitting of the monomer/oligomer equilibria according to the model shown in Fig. S3A. Resulting dissociation constants are shown in Table 1.

Detailed analysis of NmFic oligomerization. (A) SEC-MALLS elution profile (plain lines) of NmFicwt measured at varying concentrations. The eluting concentration is indicated. The corresponding apparent molecular mass values are shown overlaid (dotted lines). (B) Comparison of the structures of NmFicE156R (PDB ID code 5CKL, pink/green) and of the AD dimer of NmFicwt (dark/light gray) as part of the NmFicwt tetramer shown in Fig. 2 B and C and (C) details of the conserved interaction site. (D) Comparison of the structures of NmFicE102R (PDB ID code 5CGL, pink/orange) and of the AB dimer of NmFicwt (dark/light gray) as part of the NmFicwt tetramer shown in Fig. 2 B and C and (E) details of the conserved interaction site. (F–H) Theoretical distribution of oligomeric NmFicwt species as derived from the kinetic model shown in Fig. S3A and the Kd,1 and Kd,2 values from Table 1. (F) Distribution of the various oligomeric species of NmFicwt as a function of total concentration. All concentrations are given as monomeric molar concentrations. (G) Same as in F, but in the presence of 5 mM ATP. (H) Monomer concentration of the indicated NmFic variants as a function of total concentration.

Thermodynamic model of NmFic oligomerization. (A) Schematic view of the thermodynamic equilibrium model with associated parameters and (B) complete set of association reactions (1 to 7). D1, dimer 1; D2, dimer 2; M, monomer; Q, tetramer; T, trimer. Eq. 8 in this figure relates the change in Gibbs free energy, ∆G, upon complex formation to the dissociation constant Kd in moles per liter. Assuming no allosteric cooperativity, the ∆G associated with tetramer formation (e.g., dimerization of D2) is twice the ∆G of dimer formation (D1 in this example), because the same interface 1 is used twice. In Eq. 9 in this figure, we show that a twofold increase in ∆G results in a squared numerical value for the corresponding Kd. (C) Model of the regulatory mechanism of class III Fic proteins. Monomeric native protein (N) with unfolded αinh (N′), unfolded αinh (Y183) bound to the flap (N′′), autoadenylylated protein (A0), and autoadenylylated NmFic (A) with the modified segment detached are shown as in Fig. 6A. It is assumed that both N and A can contribute to oligomer formation, resulting in four distinct dimers (NN, 2*AN, and AA) and 16 distinct tetramers (NNNN, 4*NNNA, 6*NNAA, 4*NAAA, and AAAA), bottom part of the scheme showing the sequestration of active species. Monomeric autoadenylylated protein (A, active) binds the target protein and subsequently transfers an AMP moiety on this target. The respective binding constants (Kd,1, Kd,2, Kd,2′, and Kd,G) and kinetic parameters (kcat,1, kcat,1,eff, and kcat,2) are indicated. This model has been used to simulate the graphs shown in Fig. 6 C and D, with the thermodynamic and kinetic parameters indicated in Tables 1 and 2. This scheme remains a simplified model omitting all ATP binding steps [i.e., assuming enzyme saturation with substrate (ATP concentration >> Kd,ATP)].

To test whether the crystallographically observed tetramer also occurs in solution and whether it is of physiological relevance, we generated NmFic variants with single point mutations in one or both of the interfaces. Based on the crystal structure, charge reversal mutations E102R and E156R were expected to disrupt interfaces 1 and 2, respectively. Indeed, NmFicE102R and NmFicE156R do not form tetramers but exhibit a concentration-dependent monomer/dimer equilibrium (Fig. 2D). Furthermore, the dimer dissociation constants of the two variants agree very well with the dimer dissociation constants obtained from the fit of the NmFicwt tetramerization data (Table 1), indicating that the two interfaces are not allosterically coupled. In addition, the crystal structures of NmFicE102R and NmFicE156R (Fig. S2 B–E and Table S1) revealed virtually identical interfaces to the interfaces present in the tetrameric WT protein (Fig. S2 B–E). Finally, combination of both mutations in NmFicE102R,E156R yielded an oligomerization-deficient monomeric mutant NmFicmono, with a concentration-independent mass identical to the theoretical mass of the monomer (Fig. 2D).

Disruption of the Tetramer Activates NmFic.

To obtain insight into the physiological role of oligomerization, the tetramerization-deficient mutants were assayed in vivo. Taking a derivative of E. coli K12 MG1655 (AB472) as a model organism (8), the bacterial growth inhibition upon expression of NmFic variants was assessed on LB-agar plates supplemented with increasing amounts of isopropyl β-d-thiogalactopyranoside (IPTG; up to 2 mM) as a physiological readout for gyrase inactivation. Fig. 3A shows the quantification of bacterial growth by counting colony-forming units (CFU; also Fig. S4 A and B).

Activation of NmFic leads to E. coli growth defect and in vitro adenylylation of GyrB. (A) Quantification of E. coli growth upon repression (1% glucose) or induction of the expression of NmFic variants at varying inducer (IPTG) concentrations from a single-copy plasmid (left to right with a color gradient from white to black: 1% glucose, no IPTG, 100 μM IPTG, 500 μM IPTG, 1 mM IPTG, 2 mM IPTG). The bars represent the average of three independent experiments, and error bars represent the SD. Note the remarkable growth defect of mutants NmFicE102R, NmFicE156R, and NmFicE102R,E156R (NmFicmono). Raw data for 1% glucose and 2 mM IPTG are shown in Fig. S4 A and B. (B–D) Autoradiographs obtained after incubation of various NmFic mutants with 25 nM α-[32P]ATP, 5 mM ATP, 10 mM MgCl2, and purified GyrB43 (N-terminal GyrB 43-kDa fragment comprising the ATPase and transducer domains) at 35 °C. (B) In vitro adenylylation assay with 1 μM purified NmFic and 2.5 μM purified GyrB43 using an incubation time of 1 h. (C) Time course of autoadenylylation and GyrB43 adenylylation by NmFicE156R with or without ATP preincubation using 1 μM NmFicE156R and 5 μM GyrB43. Note that adenylylation of GyrB43 is delayed without preincubation (i.e., activation) of the Fic enzyme. (D) Same as in C, but for NmFicE156R,Y183F. Note that this mutant is unable to adenylylate GyrB43. (E–G) Quantification of the radiograph band intensities shown in B–D. The assumed error of 5% is depicted as error bars. (E) Bar diagram representing NmFic autoadenylylation (light gray) and GyrB43 adenylylation (dark gray) catalyzed by the NmFic variants. (F) Time course of GyrB43 adenylylation with or without preincubation of NmFicE156R. Also shown are the simulated curves obtained by global fitting of the kinetic model shown in Fig. 6A to the data. Resulting parameters are given in Table 2. (G) Time course of autoadenylylation of NmFicE156R and NmFicE156R,Y183F. Fitting analogously to F yielded apparent kcat,1 values of 2.4 * 10−3⋅s−1 and 4.5 * 10−4⋅s−1, respectively. AU, arbitrary units.

In vivo and in crystallo effects of residue(s) replacement in NmFic. Spotting of E. coli strain AB472 expressing NmFic derivatives on plates containing 1% glucose (repressor) (A) or 2,000 μM IPTG (inducer) (B). Note the remarkable growth defect, both in colony number and colony size of the NmFicE102R, NmFicE156R, and NmFicE102R,E156R (NmFicmono) variants. (C) Superimposition of the crystal structures of NmFicE156R (PDB ID code 5CKL, light gray) and NmFicE156R,Y183F (PDB ID code 5CMT, orange) determined at a resolution of 0.99 Å, with an rmsd of 0.23 Å for 176 Cα-positions. (D) Close-up view focused on residue 183. The 2Fo-Fc electron density map is shown in black for Y183 (NmFicE156R structure) at a contouring level of 1.2 sigma. For simplicity, only one monomer is depicted in C and D.

In contrast to WT NmFic, the tetramerization-deficient mutants NmFicE102R, NmFicE156R, and NmFicE102R,E156R showed a severe, IPTG-dependent, growth defect (Fig. 3A), resulting in the loss of almost 1.5, 3.0, and 1.5 log10 CFU/mL viability, respectively. In addition to the strongly reduced numbers, the colonies exhibited a drastically smaller size (Fig. S4B). As expected, by additional mutation of the catalytic His (H107A), the observed growth defect was completely abolished (Fig. 3A), demonstrating that the phenotype depends on the catalytic activity of NmFic. As expected, the observed E. coli growth defect correlates very well with GyrB43 adenylylation, as assayed in vitro by autoradiography (Fig. 3B). Again, the combination of the interface mutations with H107A completely suppressed both autoadenylylation and target adenylylation.

Although all tested interface mutants are active, NmFicE102R and NmFicE102R,E156R show lower activity than NmFicE156R. An explanation for this reduced activity may be that the strictly conserved acidic residue 102, which is replaced in both mutants, may contribute to the recognition of the target protein GyrB. Note that the autoadenylylation efficiency is very similar for all oligomerization-deficient mutants (Fig. 3 B and E), suggesting that the active center, per se, is unperturbed. In summary, the results indicate that NmFic tetramerization renders the enzyme incompetent for efficient autoadenylylation and for target adenylylation.

NmFic Autoadenylylates a Buried Tyr in Cis.

Automodification has been observed for most Fic proteins described to date (6, 8⇓⇓⇓⇓⇓⇓–15). For NmFic, the sites of autoadenylylation have been mapped to Y183 and Y188 of the αinh by mass spectrometry (MS) (8) (Fig. 2A, purple residues). Intriguingly, and in contrast to Y188, the strictly conserved Y183 is completely buried in the hydrophobic core of the protein. Therefore, it can be inferred that the αinh has to detach from the core of the protein for the Y183 adenylylation reaction to occur. Furthermore, for steric reasons, the αinh would no longer be able to repack after modification.

Autoadenylylation of NmFic was monitored in real-time by circular dichroism (CD) spectroscopy. Because NmFicwt shows little autoadenylylation (Fig. 3B), we used NmFicmono (NmFicE102R,E156R) for this and the following analyses. Upon addition of ATP/MgCl2, the CD spectrum of NmFicmono showed a gradual, time-dependent decrease in amplitude of the negative peaks at 208 and 222 nm (Fig. S5A), corresponding to a reduction of α-helical content. This decrease is entirely consistent with (partial) unfolding of the αinh upon autoadenylylation.

As another readout of autoadenylylation, the thermal stability of NmFicmono, was monitored by differential scanning fluorimetry (DSF; Fig. S5B). The modification resulted in significant destabilization of the protein (decrease in melting temperature of 8 °C), consistent with the partial loss of secondary structure shown above and the concomitant loss of packing interactions. Importantly, the reaction kinetics were found to be independent of the NmFic concentration, indicating that the reaction takes place in cis. The data obtained at various NmFicmono concentrations (Fig. 4) could indeed be fitted globally with one apparent first-order rate constant, kcat,1 (Table 2). Structural modeling verified that upon αinh unfolding, the disordered C-terminal segment is long enough to allow Y183 to reach the target dock of the same molecule. A model of the structural changes accompanying NmFic autoadenylylation is shown in Movie S2.

Cis-autoadenylylation of monomeric NmFic. Progress curves of autoadenylylation of NmFicmono acquired at the indicated protein concentrations using 5 mM ATP and 10 mM MgCl2. The data were fitted to a first-order reaction model. Dotted lines are the result of individual fitting, and plain lines are the result of global fitting of the three independent experiments (apparent kcat,1 of 4.5 * 10−3⋅s−1; Table 1). Note that individual and global fitting are virtually identical.

Analysis of NmFicmono autoadenylylation by real-time CD and DSF. (A) CD spectra acquired every 5 min are displayed as indicated. (Inset) Full 190- to 270-nm spectra. (B) Details of the data evaluation of the melting curves obtained by DSF. Biphasic melting curve [fluorescence (F)] of heterogeneous samples of NmFic (mixture of autoadenylylated and native protein) (Top), first derivative dF/dT (Middle), and the resulting progress curve of NmFic autoadenylylation (Bottom) are shown. Ten micromolar NmFicE156R, 2 mM ATP, and 10 mM MgCl2 were used. (C) Autoadenylylated NmFicmono (filled) and native NmFicmono (opened) concentrations are plotted as progression curves. The results of three different experiments using the same total amount of monomeric NmFic (12 μM) but distinct amounts of active (NmFicmono) and catalytically inactive (NmFicmono,H107A) monomeric NmFic are displayed as indicated. Note that only NmFicmono is autoadenylylated, but not NmFicmono,H107A, showing that autoadenylylation occurs only in cis.

Additionally, we tested whether the NmFicmono,H107A mutant, which is catalytically inactive and thus incompetent for cis modification, can be modified in trans. In mixtures of NmFicmono and NmFicmono,H107A, the final concentration of the adenylylated product was found to correspond closely to the final concentration of the active NmFicmono enzyme only (Fig. S5C). This observation corroborates the conclusion that autoadenylylation occurs in cis.

Ultimately, we subjected our samples to MS analyses (Fig. S6 A and B). Indeed, NmFicmono,H107A showed the native mass, confirming that NmFicmono is not able to catalyze the modification of the inactive mutant in trans. In contrast, the mass of NmFicmono was increased by 1,316 Da, corresponding to the mass of four AMP moieties. Thus, apart from the already identified Tyr residues Y183 and Y188, the presence of two more acceptor sites can be inferred. The MS analysis was also performed on NmFicmono,Y183F (in the presence of NmFicmono) and revealed two species corresponding to doubly and triply modified protein. As anticipated, we did not observe modification on four sites because the main acceptor site Y183 had been mutated in this protein variant. Furthermore, the Y183F mutation appears to slow down autoadenylylation (also Fig. 3G), because the automodification reaction did not reach completion (triply modified NmFicmono,Y183F). Reduced autoadenylylation of the NmFicmono,Y183F mutant compared with NmFicmono was also observed by autoradiography (Fig. S6 C and D). Finally, residues Y184 and Y185 were identified as the remaining acceptor sites, because the mutant having all four Tyr residues (Y183, Y184, Y185, and Y188) replaced by Phe showed virtually no autoadenylylation. Satisfactorily, the sequential removal of individual acceptor sites in the combinatorial mutants resulted in a linear decrease of the band intensity on the autoradiographs (Fig. S6 C and D).

Analysis of NmFicmono autoadenylylation by MS and autoradiography. (A) Mass spectrometric analysis of a 1:1 mixture of NmFicmono and NmFicmono,H107A (5 μM each) incubated for 90 min at 35 °C with 1 mM ATP and 10 mM MgCl2. NmFicmono appears fully tetra-autoadenylylated (MWtheor = 22,143 Da + 4 * 329 Da = 23,459 Da). NmFicmono,H107A shows no modification (MWtheor = 22,077 Da) confirming that the NmFic automodification does occur only in cis and not in trans. (B) Mass spectrometric analysis of a 1:1 mixture of NmFicmono,Y183F and NmFicmono,H107A (5 μM each) incubated for 90 min at 35 °C with 1 mM ATP and 10 mM MgCl2. Two species corresponding to NmFicmono being modified by two and three AMP moieties (MWtheor = 22,785 and 23,114, respectively) are observed. As in A, NmFicmono,H107A shows no modification. The experimental values of the identified species agree within 1 Da with the theoretical values. (C) In vitro target adenylylation as monitored by autoradiography. Ten micromolar purified GyrB43 was incubated with 1 μM of various purified NmFicmono variants that had one or more of the potential autoadenylylation sites on the αinh (Y183, Y184, Y185, and Y188) replaced by Phe. Incubation times were 1 h (Top) or 2.5 h (Bottom). All variants with an Y183F mutation were (largely) incompetent for target adenylylation but still showed various degrees of autoadenylylation. Only replacement of all four Tyr abolished autoadenylylation completely. It is noteworthy that all purified protein variants behaved similarly in solution. (D) Quantification of the radiograph band intensities shown in C displayed as bar diagrams after normalization (1 AU corresponding to 1 AMP moiety). GyrB43 adenylylation (Top) and NmFic auto-adenylylation (Bottom) are shown. AU, arbitrary units.

αinh of Autoadenylylated NmFic Is Partly Unfolded.

We further investigated the structural changes accompanying NmFic automodification by high-resolution nuclear magnetic resonance (NMR) spectroscopy (Fig. 5). We obtained well-dispersed NMR spectra for an unmodified protein (catalytically inactive NmFicmono,H107A), as well as for autoadenylylated (NmFicmono-AMP) protein. Sequence-specific backbone resonance assignments were obtained for 96% of the residues of NmFicmono,H107A (Fig. S7A). Based on these assignments, the chemical shifts of the autoadenylylated form could be assigned to 91% and confirmed by additional triple-resonance experiments. The overlay of the 2D [15N,1H]-HSQC (heteronuclear single quantum coherence) spectra of the native and autoadenylylated proteins shows large chemical shift differences for a subset of resonances (Fig. 5A). The residue-specific analysis of chemical shift changes upon autoadenylylation (Fig. 5 B and C) revealed that helix 8 (αinh) is the most affected region of the protein. Therein, the covalently adenylylated residue Y183 showed the most pronounced chemical shift change [Δδ(HN) = 6.7 ppm]. Furthermore, the amide chemical shifts of residues 178–191 populate the random coil region (7.5–8.5 ppm), indicating an unfolded conformation. In addition, the target-binding site (flap) and helix 1 (adjacent to helix 8) show significant chemical shift differences, but to a lesser extent than the αinh, and a few residues in these regions are broadened beyond detection (Fig. 5B). Due to the point mutation H107A in the native form, the active site region also shows some chemical shift differences (Fig. 5 B and C). Despite these changes, the structural scaffold of the protein remains intact upon autoadenylylation. To characterize the secondary structure elements of both NmFic forms in solution, we used the secondary chemical shifts of the backbone 13Cα nuclei (Fig. 5D). In aqueous solution, NmFicmono features eight α-helices, which agree in number and positioning with the crystal structure. The picture changes for the autoadenylylated form, in which the first seven helices are maintained, whereas the αinh features helical structure only in its first two helical turns, but is unfolded from the position of residue G178 onward. These secondary structure changes are in full agreement with the chemical shift perturbation in 2D [15N,1H]-HSQC spectra (Fig. 5D and Movie S2). Overall, these data confirm the hypothesis that the αinh cannot adopt its original position and conformation upon autoadenylylation of Y183.

NmFic Autoadenylylation Relieves Autoinhibition.

We have shown that cis-autoadenylylation of NmFic constitutes an in-built mechanism to covalently modify Y183, resulting in partial unfolding of the αinh. Obviously, this modification should be of functional relevance, considering that the αinh in its native form partially obstructs the nucleotide binding site, and thereby autoinhibits Fic enzymes (8). Thus, to test for any functional role of Y183, we removed its hydroxyl group by introducing an additional Y183F mutation into the alleles of active (i.e., oligomerization-deficient), mutants. Strikingly, the mutation suppressed the growth defect phenotype as efficiently as mutation of the catalytic His (H107A) (Fig. 3A, Right), suggesting a crucial regulatory role for Y183.

The in vivo observation was faithfully mirrored in the autoradiograph obtained after incubation of the various purified NmFic mutants with radioactive ATP (Fig. 3B). In vitro GyrB43 adenylylation, which is efficiently catalyzed by the three interface mutants (Fig. 3B, lanes 4–6), is almost completely abolished in their respective Y183F variants (Fig. 3B, lanes 10–12).

The remaining autoadenylylation of the Y183F NmFic variants (Fig. 3B, Right), which is due to the modification of Y184, Y185, and Y188, indicates that enzyme function, per se, is not impaired. As expected, no target adenylylation is observed. Thus, (partial) modification of the additional Tyr residues does not significantly relieve autoinhibition. Still, to corroborate that the Y183F mutation had no unforeseen effect on the enzyme, we determined the high-resolution (0.99 Å) crystal structures of both NmFicE156R and NmFicE156R,Y183F (Fig. S4 C and D). Indeed, the structures are virtually identical, apart from the absence of the Y183 hydroxyl group in the double mutant.

To test whether the kinetics of autoadenylylation and target adenylylation would be of physiological relevance, we measured time courses of product (GyrB-AMP) formation for NmFicE156R (Fig. 3C). The apparent rate of NmFicE156R autoadenylylation derived from Fig. 3G agrees well with the one obtained for NmFicmono by DSF analysis (Table 2). Similarly, NmFicE156R,Y183F shows autoadenylylation, although with a considerably slower rate (Fig. 3 D and G). Such an effect of the Y183F mutation was also observed for NmFicmono,Y183F by MS (Fig. S6B). As mentioned above, the modification on residues Y184, Y185, and Y188 is probably not of functional relevance.

Under the used conditions, NmFicE156R fully converts GyrB43 to its adenylylated form within 1 h (Fig. 3 C and F), but with a pronounced lag phase. This delay is absent in the progress curve of preactivated Fic enzyme, which suggests again that only the adenylylated form is catalytically active. This interpretation was confirmed quantitatively by globally fitting the respective kinetic model (Fig. 6A, but without oligomerization) to the data (Fig. 3F). Upon setting Km,GyrB to the experimentally determined value of 39 μM, the fit yielded the turnover numbers of autoadenylylation (kcat,1,eff) and target adenylylation (kcat,2) given in Table 2. Notably, kcat,1,eff turned out to be about one order of magnitude slower that the corresponding rate, kcat,1, as measured by DSF. Therefore, it can be concluded that the rate-determining step of Fic activation is the debinding of the modified segment from the active site (A0→A).

Concentration and time dependence of NmFicwt-catalyzed target adenylylation. (A) Simplified model of the regulatory mechanism of class III Fic proteins. (Top Left) Native monomeric Fic protein (N) is activated by autoadenylylation to yield A. This reaction involves unfolding of the αinh (N′), binding of the modifiable Tyr Y183 of the unfolded segment to the flap (N′′), and subsequent autoadenylylation of Y183 (A0). (Bottom) Both types of monomers (N and A) are in dynamic monomer/tetramer equilibrium to form homotetramers or heterotetramers NiAj, with i + j = 4. The tetramer acts as a buffer sequestering the active monomers. (Right) Only the remaining fraction of free (monomeric) A molecules is competent for target adenylylation. The complete model with all thermodynamic and kinetic parameters is shown in Fig. S3C. (B) Autoradiographs showing NmFicwt and GyrB43 adenylylation. The same experimental setup as in Fig. 3C was used but was carried out at various NmFicwt concentrations (1 nM to 2 μM, as indicated). Reactions were stopped after the indicated incubation times. (C) Simulation of GyrB adenylylation as a function of total NmFicwt concentration based on the kinetic model shown in A and Fig. S3C, with parameters set to their measured values (Tables 1 and 2). (D) Replot of the data shown in C as a function of incubation time for representative NmFicwt concentrations.

Activity Profile of NmFicwt Is a Consequence of Autoactivation Combined with Oligomerization.

We have thus shown that NmFic-catalyzed target adenylylation is controlled by enzyme tetramerization as well as cis-autoadenylylation. These results were obtained by analyses of mutants deficient in one or the other function. What, then, would be the combined effect in the WT enzyme? Fig. 6B shows the autoradiographs obtained after incubation of NmFicwt (at varying concentrations) with GyrB43 for various durations. At the standard enzyme concentration of 1 μM, weak NmFicwt autoadenylylation and no significant target adenylylation are observed for incubation times up to 8 h, consistent with our earlier result (Fig. 3B, first lane). Strikingly, strong GyrB43 adenylylation is observed at lower enzyme concentrations (250 nM and lower), with an abrupt transition from 250 to 500 nM. Because this transition occurs in a very similar range as the monomer-to-tetramer transition (Fig. 2D, Bottom), it most probably reflects the emergence of catalytically incompetent tetramers at higher concentrations.

Importantly, however, the catalytic incompetence of the tetramer alone is not sufficient to explain the data, because not only the tetramer/monomer ratio but also the absolute concentration of active monomers will increase with total concentration (Fig. S2H). To rationalize the observed effects quantitatively, we set up a kinetic scheme including autoactivation and inactivation by oligomerization as shown in Fig. 6A (also Fig. S3C). In this model, autoinhibited native NmFic (N) is in equilibrium with states N′ (αinh unfolded) and N′′ (modifiable Y183 positioned in the active site). The latter state gets autoadenylylated at Y183 in a first-order reaction to yield A0. Finally, unbinding of the modified C terminus will result in a monomer state A that is predicted to be competent for target adenylylation. Note, that ATP binding will shift the A0⇔A equilibrium toward the competent A state. Because monomers are in fast oligomerization equilibrium, active A monomers will repartition into the tetramer that acts as a reservoir, and will therefore be partly sequestered to the inactive oligomeric state.

We reasoned that a contribution of the unfolded autoadenylylated C-terminal segment to the interface might affect the oligomerization affinity. Indeed, in presence of 5 mM ATP, the dissociation constant (Kd,2′) of A was measured by AUC-SV to be fivefold lower than the corresponding constant of N (Fig. 2D and Table 1).

Using the experimentally determined parameters of the kinetic model (Tables 1 and 2), we then simulated GyrB43-AMP production as function of enzyme concentration and incubation time by numeric integration of the respective differential equations (Fig. 6 C and D). Indeed, as observed experimentally, target adenylylation efficiency drops with enzyme concentration above a certain threshold (Fig. 6C). Furthermore, the simulated progress curves (Fig. 6D) reproduce the lag phase observed at a high enzyme concentration. Taken together, the salient features of intrinsic NmFic regulation are faithfully captured by the proposed kinetic model.

Discussion

The physiological importance of AMP-transferases with FIC fold has escaped attention for a long time, because their activity is kept in check by intra- or intermolecular active site obstruction (8). In this study, we focused on class III Fic proteins that carry the inhibitory αinh at the C terminus and are composed of a FIC domain only. We reveal that in addition to the previously reported autoinhibition (8), two further mechanisms inversely regulate adenylylation activity, namely, tetramerization and cis-autoadenylylation. This combination results in a complex autoregulatory mechanism.

NmFic forms a tetramer with the involved interfaces largely conserved or coevolved among class III members, suggesting conservation of the tetrameric arrangement in this class. Disruption of either interface was achieved by site-directed mutagenesis, thus verifying their role both in the crystalline state and the solubilized state. Due to the cooperative nature of tetramerization, a sharp monomer–tetramer transition occurs in the presence of ATP, at the rather low NmFic concentration of 50 nM (Fig. 2D and Fig. S2G). Clearly, the tetramer state is of physiological relevance, because expression of interface disruption mutants, but not of NmFicwt, resulted in impaired growth of the expressing E. coli strain. Thus, it can be inferred that only monomeric NmFic is able to exert the growth retardation effect, which can be attributed to GyrB adenylylation (5). Most probably, the tetramer is catalytically noncompetent, because the binding site for the segment flanking the modifiable side chain [target dock (10)] is partially occluded. Furthermore, the strict conservation of oligomerization interface 1 (as opposed to interface 2, which shows covariation) may point to its involvement in mediating the contact between (monomeric) NmFic and the target. In fact, the surface-exposed area of the tetramer is highly variable, and therefore probably not involved in target recognition (Movie S1).

Fic automodification has been reported repeatedly (6, 8⇓⇓⇓⇓⇓⇓–15), but its role has remained unclear. In fact, nonspecific modification due to the high in vitro Fic concentrations used seemed possible. Here, we have shown that NmFic autoadenylylation crucially controls enzyme activity by relieving the autoinhibitory effect of the αinh, as demonstrated by the Tyr-to-Phe mutation (Y183F) that fully suppressed the activating effect of the interface disruption mutants (Fig. 3A) and rendered the enzyme incompetent for target adenylylation (Fig. 3B). Biochemical and structural analyses confirmed that the conservative Y183F mutation does not disrupt the FIC fold (Fig. 3 and Fig. S4 C and D).

Importantly, the automodification occurs in cis with a reaction velocity independent of the total enzyme concentration. Indeed, structural modeling shows that upon unfolding of the αinh, the modifiable Tyr can reach the active site of the same molecule (Movie S2). Probably, the segment flanking the Tyr residue would engage in β-strand interaction with the flap, resulting in correct registration of the modifiable tyrosyl side-chain within the active site. This binding mode would be analogous to the one seen in the complex structures of IbpAFic2/cdc42 (6) and of noncognate peptide/Fic proteins (9). Because the flap is partially buried at the center of the tetramer, the structural model also explains why the tetramer would be incompetent for automodification. Thus, in the presence of ATP, native monomeric NmFic gets efficiently converted to its active, covalently modified form with a rather fast “in-built” first-order rate constant (Table 2, equivalent to a t1/2 of about 150 s). As such, NmFic and, most likely, Fic proteins of class III in general constitute self-contained proteinaceous timers. Whether such an autoactivation mechanism also pertains to Fic proteins of class I and II remains to be seen, but conserved Tyr acceptor candidates can be identified in the N-terminal and central part of the fold, respectively. Noteworthy, cis-automodification has also been described for a few kinases as an activation mechanism (18).

The combination of the two coupled and counteracting effects, automodification and tetramerization, results in a quite unusual time and concentration dependence of target adenylylation, as captured in vitro by autoradiography (Fig. 6B) and reproduced in silico by respective simulations (Fig. 6 C and D). In essence (Fig. 6A), monomeric native Fic molecules (N) get converted with the intrinsic first-order rate to the autoadenylylated, active form (A). However, because Fic monomers are in fast equilibrium with the tetrameric state, they get redistributed such that the A/N ratio will be equal or larger in the tetrameric state. As a consequence, the increase in the absolute number of active monomers (A) will be dampened. Obviously, this buffering effect is most pronounced at high tetramer/monomer ratios (i.e., high enzyme concentrations) and will diminish when the autoadenylylation reaction reaches completion.

The comprehensive characterization of the complex autoregulatory mechanism of class III Fic proteins presented here sets the stage for future investigations of its physiological consequences. It has to be inferred, however, that the intrinsic regulatory mechanism may be further modulated by external factors, such as antagonistic phosphodiesterases [e.g., the de-AMPylase SidD of Legionella (19, 20)], specific proteases, or small ligands. Furthermore, Fic biosynthesis and degradation, as well as dilution effects during cell growth and division, will profoundly affect the time profile of the Fic pool activity. Still, solely based on the inherent mechanism, interesting consequences can be envisaged.

The nonmonotonic dependence of AMP-transferase activity on enzyme concentration would render the enzyme exquisitely sensitive to variation of its concentration in the bacterial cell. A twofold drop of NmFic concentration in the right range (e.g., due to cellular growth and division) would drastically increase catalytic activity, and thus GyrB inactivation. In addition, the considerable delay in NmFic-mediated GyrB adenylylation may be important in vivo to ensure that the growth-retarding effect of the target modification kicks in only under certain conditions, such as starvation, when synthesis of fresh NmFic has come to a halt. Molecular timers have been found before, such as the master regulator Spo0A of Bacillus that controls, upon phosphorylation, the switch from competence to sporulation via a well-characterized genetic circuit (21, 22) induced by prolonged times of nutriment starvation. Class III Fic proteins may constitute a new family of molecular timers that are, in contrast to the master regulator Spo0A of Bacillus, fully autonomous and not relying on any feedback exerted by other components or a genetic circuit. This hypothesis will have to be tested in future studies.

Materials and Methods

Detailed information on cloning, expression, purification, crystallization, data collection, and structure determination by X-ray crystallography or NMR is provided in SI Materials and Methods. Plasmids were constructed as described previously (8, 23, 24) (Tables S2 and S3). Proteins were expressed and purified as described (8, 25). Toxicity tests and adenylylation assays were performed according to the protocols given by Harms et al. (5) and Goepfert et al. (9).

The oligomeric state of NmFic was determined by SEC-MALLS and AUC-SV at varying concentrations. DSF (26) was used to monitor in vitro autoadenylylation of NmFic. For simulations and fitting of functional data to the kinetic model, the programs ProFit 6.2.14 (QuantumSoft) and Complex Pathway Simulator (COPASI) 4.14 (27) were used.

SI Materials and Methods

Plasmid Construction.

pRSF-Duet1 derivatives were cloned as described previously (8) and used for overexpression of proteins for purification. For phenotypic analyses, nmfic genes were cloned into the single-copy vector pNDM220 (23) under the control of a particularly tight Plac. Site-directed mutagenesis was performed following the protocol described by Zheng et al. (24). A list of used plasmids and a list of respective oligonucleotides can be found in Tables S2 and S3.

In Vitro Adenylylation Assay.

Adenylylation assays were performed using cleared cell lysate of ectopically expressing E. coli as described previously (8) [except using BL21 instead of BL21 (λDE3) cells for overexpression] or using purified proteins as described by Goepfert et al. (9). Adenylylation activity of NmFic was assessed by incubating NmFic with 5 mM ATP, 10 μCi [α-32P]-ATP (Hartmann Analytic), 25 mM MgCl2, and GyrB43. In addition, 200 μM novobiocin was added to inhibit the ATPase activity of GyrB43. Adenylylation assays with purified NmFic and GyrB43 were performed at 35 °C, and adenylylation assays using cell lysates were performed at 30 °C. For data evaluation, the intensity of each band on the autoradiographs was quantified using ImageJ (rsbweb.nih.gov/ij/download.html).

Structure Determination.

X-ray data were collected at the Swiss Light Source (Villigen, Switzerland) on beamline X06SA (PXI) or X06DA (PXIII) at 100 K (Table S1). Diffraction data were indexed and integrated using XDS (32) and subsequently merged and scaled using XSCALE (32) or AIMLESS (33). Data collection and processing statistics are given in Table S1. Structures were determined by molecular replacement [PHASER program (34)] with an NmFicwt monomer [PDB ID code 3S6A (8)] as a search model. Several rounds of model building and refinement were performed using Coot (35) and REFMAC5 (36) or phenix.refine (37). Five percent of the data were excluded from refinement and used for cross-validation. The geometry of the final model was assessed using MolProbity (38) showing >99% of the residues in the core and allowed regions of the Ramachandran plot. Final refinement yielded NmFic models with reasonable Rwork/Rfree values for their respective resolution ranges. Refinement statistics are summarized in Table S1. Figures were prepared with DINO (A. Philippsen; www.dino3d.org).

Conservation and Covariance Analyses of Class III Fic Proteins.

Conservation scores for each position of the class III Fic protein NmFic were calculated by using the ConSurf server with default parameters (39). The top 200 sequences (46% identity) were selected from a BLAST search against the Uniref90 database and manually curated to select only class III Fic proteins (three sequences that do not contain the C-terminal αinh were removed). For covariance analysis, the NmFic sequence was submitted to the Gremlin server for coevolution analysis (29).

NMR Spectroscopy.

NMR experiments were performed on Bruker 600-, 700-, and 900-MHz spectrometers running Topspin3.0 and equipped with cryogenically cooled triple-resonance probes. All experiments were performed in NMR buffer [25 mM MES, 150 mM NaCl (pH 6.5)] at 25 °C. For the sequence-specific backbone resonance assignments of NmFic, the following experiments were recorded: 2D [15N,1H]-TROSY-HSQC (40), 3D HNCA (41), 3D HNCACB (41), 3D HNCO (41), and 3D HN(CA)CO (41). To confirm the chemical shift changes observed upon adenylylation within 2D [15N,1H]-TROSY-HSQC, the following 3D experiments were recorded in a nonuniformly sampled manner with a random sampling schedule generated by Topspin3.0 (Bruker Biospin) to confirm the assignments: 3D HNCA (25% of the full time-domain grid) and 3D HNCO (9%).

For the analysis of the dynamic properties of NmFic, the following experiments were measured: 15N{1H}-NOE (42), T1(15N) (42), and TROSY for rotational correlation times (TRACT) (43). NMR data were processed with PROSA (44), NMRPipe (45), and mddNMR2.4 (46), and were analyzed with CARA, XEASY (47), and Topspin3.0 (Bruker Biospin). Nonlinear least square fits of relaxation data were done with MATLAB (MathWorks). R2(15N) values were derived from Rα(15N) and Rβ(15N). Error bars for R1(15N), Rα(15N), and Rβ(15N) were calculated by a statistical bootstrapping scheme, and error bars for the 15N{1H}-NOE were calculated from the spectral noise. Secondary chemical shifts were calculated relative to the random coil values of Kjaergaard and Poulsen (48).

The sequence-specific resonance assignment for the apo NmFic has been submitted to the Biological Magnetic Resonance Data Bank under accession code 26607. The chemical shift changes of the amide moiety upon ATP binding were fitted by nonlinear regression analysis to Eq. S1 by using standard software. The term ∆obs corresponds to the chemical shift difference at a given titration point, and ∆max is the maximal chemical shift difference at the last titration point:Δobs=Δmax(Kd+[ATP]0+[NmFic]0)−(Kd+[ATP]0+[NmFic]0)2−(4[NmFic]0[ATP]0)2[NmFic]0.[S1]

SEC-MALLS Analysis.

SEC-MALLS was used for the determination of the NmFic oligomerization dissociation constants by measuring apparent mass values (MWapp) at various loading concentrations. UV absorption and differential Refractive Index (dRI) values were used to derive the eluting concentration. A similar experimental setup has been described previously; details are provided in the study by Sundriyal et al. (49). The column was preequilibrated with 10 mM Tris (pH 7.6) and 100 mM NaCl at 4 °C.

Based on the respective mass action law equations (Fig. S3B), the dependence of MWapp as a function of protein concentration was modeled. Assuming that both dimer interfaces (Fig. 1) are independent, the thermodynamic tetramerization scheme (Fig. 3A) contains only two independent parameters, the dissociation constants of the respective interfaces (Kd,1 and Kd,2). For data fitting, the reaction extents were used as state variables and inequality constraints were imposed to ensure the correctness of the results. Optimization of reaction extent values was accomplished using the sequential least squares programming method (50) as implemented in the SciPy library (51). For mutants NmFicE102R and NmFicE156R, the experimental data were fitted to a simple monomer/dimer equilibrium. All equilibria were modeled as fast-exchange processes with respect to the time scale of chromatographic separation, because all measurements yielded only a single MALLS peak. Theoretical MWapp values were calculated from mass concentrations (cm,i) and molecular weights (MWi) of populated species (monomer and dimer for the NmFicE102R and NmFicE156R mutants; monomer, dimer, trimer, and tetramer for NmFicwt) asMWapp=∑icm,iMWi∑icm,i.[S2]All routines were implemented in the Python language using NumPy and SciPy numerical libraries (51).

AUC Analysis.

NmFicwt (40 μM) was labeled with Dylight 488 NHS ester using the protocol published by the supplier (Thermo Scientific). A dilution series of unlabeled NmFicwt (10.0 μM, 3.33 μM, 1.11 μM, 0.370 μM, 0.123 μM, 41.0 nM, and 0 nM) supplemented with 40 nM fluorescein-labeled NmFicwt was prepared in the presence and absence of 5 mM ATP. SV experiments were performed on 400-μL samples in double-sector charcoal-Epon centerpieces at 42,000 rpm and 6 °C using a Beckman XL-I analytical ultracentrifuge with the Beckman An-50 Ti rotor. Sedimentation was monitored in 1,000 scans during an overnight run using the Aviv AU-FDS fluorescence detection system. The buffer density (1.0022 g/mL) and viscosity (0.01474 poise) were measured at 6 °C using an Anton Paar DMA 4500M density meter and AMVn viscometer. Similarly, a dilution series of unlabeled autoadenylylated NmFicwt-AMP (1.25 μM, 625 nM, 312.5 nM, 156 nM, 78 nM, and 0 nM) supplemented with 40 nM fluorescein-labeled autoadenylylated NmFicwt-AMP was prepared in the presence of 5 mM ATP. SV experiments were performed using a Beckman XL-I analytical ultracentrifuge with the An-60 Ti rotor. The SV data were fitted to a diffusion-deconvoluted sedimentation coefficient distribution [c(s)] using the software Sedfit (52). A sedimentation coefficient range of 1–15 S was used with a resolution of 200 points. Radially-invariant (RI) noise and time-invariant (TI) noise, baseline, and meniscus position were all fitted. In a first round of fitting, the frictional ratio f/f0 was fitted for each sample. The data were then refitted using an average value for f/f0 (1.11) calculated from the free-fitted values. The c(s) distributions were overlaid and normalized by maximum signal intensity in the software GUSSI (Chad Brautigam, University of Texas Southwestern Medical Center, Dallas, TX), and weight-averaged sw isotherms were calculated from the c(s) distributions over the range 1–5 S. The fraction of monomeric (xM) and tetrameric (xQ) NmFic species (with sedimentation coefficients of 1.62 S and 3.95 S, respectively) as a function of protein concentration were derived from the relative areas of the respective peaks. Finally, the xM and xQ distributions were fitted to the kinetic scheme to yield the dimerization constants (as discussed above).

CD Analysis.

A time series of CD spectra was recorded at 35 °C in a quartz-suprasil cuvette with a path length of 0.2 cm using the wavelength range 190–270 nm (26.67 nm⋅min−1) on an Applied Photophysics Chirascan Plus CD spectrophotometer. A measurement was recorded every 5 min. The autoadenylylation reaction was started in the cuvette (t0) by addition of 1 mM MgCl2 to a solution containing 20 μM NmFicmono, 0.5 mM ATP in 10 mM Tris (pH 7.8), and 125 mM NaCl. The α-helical content was approximated (52) by dividing the mean residue ellipticity at 222 nm by −30,000 deg⋅cm2⋅dmol−1.

DSF.

Autoadenylylation was followed in vitro by dye-based thermal denaturation experiments (26) performed on a Qiagen Rotor-gene Q real-time PCR cycler. Experiments were carried out in 200-μL thin-wall PCR tubes (Thermo Scientific) with 50 μL of sample containing the indicated concentration of NmFic and ATP, 10 mM MgCl2, and 5× SyPro Orange dye (Invitrogen) in 10 mM Tris (pH 7.8) and 100 mM NaCl, except when stated differently. For time-course measurements, reactions were stopped at the indicated times by addition of 50 mM EDTA to an aliquot of the reaction. Temperature was increased by a 0.5 °C increment every 30 s between 20 °C and 90 °C. Fluorescence intensity (F) thermal denaturation curves and their first derivatives (dF/dT) were generated using Rotorgene software (Qiagen) and exported as text files. The first derivatives were fitted as the sum of Gaussian peaks using ProFit (QuantumSoft). The fraction of native and autoadenylylated NmFic was derived from the relative areas of the fitted peaks.

Kinetic Fitting and Simulations.

Enzymatic reactions were fitted to standard first-order and second-order reaction kinetics combined with dynamic protein oligomerization (Fig. 6A), which yielded the apparent rate constants kcat,1 and kcat,2 for autoadenylylation and target adenylylation, respectively, at the given substrate (ATP) concentration. The NmFic-GyrB43 dissociation constant (Kd,G) was set equal to the Km value obtained from GyrB titration. Oligomerization dissociation constants (Kd,1, Kd,2, and Kd,2′) were set to their experimentally determined values. From the fit of NmFicE156R-catalyzed GyrB adenylylation, the effective autoadenylylation rate (kcat,1,eff) was obtained. Numerical integration of the differential equations corresponding to each reaction (described in the legend for Fig. S3C) was performed using the Complex Pathway Simulator (COPASI) software (27). Fitting and simulations were performed using either ProFit 6.2.14 (QuantumSoft) or COPASI 4.14.

Acknowledgments

We acknowledge Gerd Pluschke (Swiss Tropical and Public Health Institute) for kindly providing the genomic DNA of N. meningitidis 2808. We thank Tillmann Heinisch for help with MS analysis. We thank the staff of beamlines X06DA and X06SA of the Swiss Light Source for excellent support. This work was supported by the European Research Council Advanced Investigator Grant (ERC-2013-AdG) FICModFun 340330 (to C.D.) and Swiss National Science Foundation Grants 3100-132979 (to C.D.) and 31003A-138414 (to T. Schirmer).

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